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Department of Pathology and Laboratory Medicine, James Hogg iCAPTURE Centre for Cardiovascular and Pulmonary Research, St. Paul's Hospital/Providence Health Care, University of British Columbia, Vancouver, British Columbia, Canada V6Z 1Y6
Submitted 29 August 2003 ; accepted in final form 26 January 2004
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ABSTRACT |
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length adaptation; oscillation; myosin filament; actin cytoskeleton; morphometrics
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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11 x 1 x 0.4 mm in dimension. The strips of muscle were affixed to aluminum foil clips at both ends and mounted in a muscle bath. One end of the strip was joined to a stationary hook and the other end to a length-force transducer with a signal-to-noise ratio >50 and a compliance of 1 µm/mN (QJin Design, Winnipeg, MB, Canada). The apparatus and tissue preparation have been described previously (33, 34). The muscle bath contained PSS with pH 7.4 at 37°C and bubbled with a gas mixture (5% CO2-95% O2). The PSS has a composition (in mM) of 118 NaCl, 4.5 KCl, 1.2 NaH2PO4, 22.5 NaHCO3, 2 MgSO4, and 2 CaCl2 and 2 g/l dextrose.
Experimental procedures.
Before a trachealis preparation was chemically fixed for electronmicroscopic examination, it was equilibrated at a preset length for 1 h. During the equilibration period, the muscle was stimulated (with electric field stimulation, 60 Hz) periodically to produce 12-s tetani at 5-min intervals. The preparation was considered equilibrated when it developed a stable maximal isometric tetanic force. There were five experimental conditions under which the preparations were fixed (after equilibration): 1) relaxed and unstrained (at the in situ length), 2) activated at the in situ length, 3) relaxed and strained (at 1.5 times the in situ length), and 4) activated and strained (at 1.5 times the in situ length), and 5) relaxed at the in situ length and postoscillation (the length-oscillation frequency was 0.5 Hz; the peak-to-peak amplitude of strain was 60% of the in situ length; the oscillation was centered around the in situ length, which meant that the cells were subjected to a 30% stretch beyond their in situ length; and the duration of oscillation was 5 min). For the preparations fixed under an activated state, the final stimulation was produced by addition to the muscle bath 0.1 mM of acetylcholine; the muscle was fixed at the plateau of isometric contraction 120 s after the addition of acetylcholine. The average stress was found to be 101.6 ± 11.6 (SE) kPa. Acetylcholine (instead of electric field stimulation) was used in the final contraction to ensure that activation of the muscle was maintained during fixation.
Electron microscopy.
Tracheas from three animals were used for the electron microscopy analysis. Five strips of muscle per trachea were dissected and equilibrated at the preset lengths according to the five experiment conditions described above. Muscle preparations were fixed for electron microscopy by using a conventional protocol described previously (18, 27). Briefly, muscle preparations were fixed with the primary fixing solution (see below for details) for 15 min while they were still attached to the experimental apparatus where isometric force was continuously monitored. The tissue was then removed from the apparatus and cut into small cubes and immersed in the primary fixing solution for an additional 2 h at 4°C. The primary fixing solution contained 2% glutaraldehyde, 2% paraformaldehyde, and 2% tannic acid in 0.1 M sodium cacodylate buffer. In the process of secondary fixation, the cubes were put in 1% OsO4 in 0.1 M sodium cacodylate buffer for 2 h. The tissue was then stained with 1% uranyl acetate, dehydrated with increasing concentrations of ethanol, and embedded in resin (TAAB 812 mix). The blocks were sectioned with a diamond knife to obtain sections of 90 nm of thickness. The sections (on copper grids) were further stained with 1% uranyl acetate and Reynold's lead citrate. Images of longitudinal and cross-sectional areas of smooth muscle cells were obtained by using a Phillips 300 electron microscope.
Morphometric analysis.
Sampling and analysis were carried out "blind." The codes indicating experimental conditions were revealed only after the analysis of each group was finished. A total of 25 electron micrographs of muscle cell cross sections per trachea were analyzed (5 per preparation). A specialized image-analysis software (Image Pro-Plus 3.0) was used to help in the manual counting of the thin filaments by marking and keeping track of the number of filaments counted (tag-point counting). The computer program randomly placed 10 circles with an area of 0.1734 µm2 on each micrograph. Circles that fell on the areas occupied by nucleus, organelles, plasma membrane, or extracellular space were excluded from the study; only the circles that fell on cytoplasmic areas with mostly thick and thin filaments, and occasionally dense bodies, were counted. The area occupied by dense bodies was not included in the calculation of filament density. Of the 10 circles generated by the computer, 3–4 of them met the criteria. The thin filaments were counted by the tag-point quantification method in each of the nonexcluded circles placed in the cytoplasmic areas on the cells. According to standard morphological methods, the filaments that fell on the edge of the circles were counted as half. The total numbers of thin filaments counted were divided by the area of the circle (minus area of dense bodies, if there were any) to obtain the thin filament density. For average thick filament density, because the number of thick filaments per cross section is relatively small, the total number of thick filaments within a cell cross section was obtained and divided by the cytoplasmic area of that cell cross section.
Statistical analysis. The analysis and comparison between the groups were performed by one-way or two-way ANOVA. The n value was the number of animals used. Data from each animal (5–6 micrographs per animal) were averaged first before the means from different animals were averaged. Statistically significant difference corresponds to a P value of <0.05.
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RESULTS |
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6–7 nm. On average, 3–4 circles (out of the 10 randomly generated by the computer) per micrograph met the selection criteria and were analyzed.
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15% of the area within the circle) selected for examination for thin filament density around the dense bodies. The central and peripheral thin filament densities were assessed in the same way as described for thick filament density assessments (described above for Fig. 4). The filament density was examined under four conditions: relaxed vs. contracted at 1.0 times and 1.5 times the in situ length. For the comparisons of densities near the dense bodies, six micrographs from each group were used, and in each micrograph thin filaments in three circles were counted. The percentages of areas occupied by the dense bodies within the circles for the relaxed and contracted groups were 16.1 ± 0.1 and 14.8 ± 0.1%, respectively. They were not statistically different (P = 0.3). For the comparisons of central and peripheral thin filament densities, six micrographs from each group were used, and in each micrograph thin filaments within one circle placed in the central region of the cell cross section and one circle placed in the peripheral region were counted. Figure 6 summarizes the comparisons. Thin filaments were more concentrated near the dense bodies; they increased by 11.6 ± 2.8 (SE) and 11.5 ± 3.9% in the relaxed and activated cells, respectively. On activation, the density near a dense body increased by 19.2 ± 1.9% (SE), nearly identical to the increase in density in the randomly selected area during activation (19.5 ± 3.9%). Two-way ANOVA revealed that the elevated thin filament densities around dense bodies and after contractile activation were significant (P < 0.05), and there was no statistical interaction between dense body area-specific and contractile state-associated thin filament densities. The distribution of thin filaments in a cell cross section appeared to be even, and there was no systematic aggregation of the filaments in the central or peripheral areas after the cells had been activated or stretched (Fig. 6).
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DISCUSSION |
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Before we discuss the results in detail, it is important to point out limitations of our measurements. There was a small change in the angle of alignment of contractile filaments with the cell's longitudinal axis during contractile activation (as the filaments became more parallel to the axis of force transmission, Fig. 2); this could affect the filament density count in cell cross sections. However, the fact that the angles were in general relatively small is not likely to change the conclusion of this study (although it may quantitatively alter the density values). The filament density measured in a cell cross section at different cell lengths is directly proportional to the mass of the filaments in the cell, if the cell volume is constant. Because the filament mass is determined by the average length and number of the filaments, the filament density (D) is a product of filament length (L) and number (N); i.e., D = L x N. Unfortunately, we cannot differentiate a density change due to a change in filament length from that due to a change in the number of filaments. An increase in filament density can only be interpreted as a result of three possible changes: 1) an increase in the number of filaments, 2) an increase in filament length (of existing filaments), or 3) a mix of the above two possibilities.
Activation-dependent thin filament density and its regional variation. Actin polymerization during contractile activation has been examined by many investigators using a variety of methods; pharmacological inhibition of the transition from globular (G)- to filamentous (F)-actin has been extensively used in the studies of both intact and cultured smooth muscle cells (1, 7, 20, 22, 25, 28); immunostaining of both G- and F-actins has also been used (4, 8, 14–15); biochemical quantification is another method used to assess the composition of G- and F-actins (23, 30). The above-mentioned studies examined the "global" changes in the contents of G- and F-actins in muscle cells; the methods used in these studies lacked the resolution to pinpoint where within the cytoplasm the polymerization or depolymerization of actin occurs in response to specific stimuli. The results presented in the present study revealed for the first time the difference in actin filament densities in the subcellular domains. The thin filament density near the cytoplasmic dense bodies was found to be slightly higher than the density in the cytoplasmic areas without dense bodies (Fig. 6). This small but significant increase in density suggests that there was a tendency for thin filaments to bundle together near dense bodies; this is perhaps related to the appearance of "myofibrils" within individual smooth muscle cells found in longitudinal sections where thin filaments attach to dense bodies in a horsetail fashion (3). The densities in the central and peripheral areas of cell cross sections were, however, not altered by contractile activation or stretching of the cells (Fig. 6).
Although a significant increase in thin filament density was found to accompany contractile activation (Fig. 3), there was no out-of-proportion regional increase (Fig. 6). The present finding is consistent with the interpretation that the actin polymerization is more or less even globally, not localized to any particular subcellular domains in a cell cross section. Polymerization of myosin thick filaments due to contractile activation, on the other hand, is rather inhomogeneous (Fig. 1B), as also reported by our laboratory previously (13, 16, 17). The clustered aggregation of thick filaments in activated cells, however, does not show systematic concentration of thick filaments in the central region of the cell or vice versa (Fig. 4).
It is still not clear why polymerization of actin is needed in smooth muscle during activation. There appears to be abundant thin filaments packed within the cytosol, even in the relaxed state. It has been postulated that lengthening of thin filaments might be needed for focal adhesion (9). The nonlocalized thin filament lengthening and formation observed in the present study suggest that the polymerization may have a function of facilitating thin-thick filament interaction and therefore force generation. However, it would appear that the relatively sparse thick filaments would be the limiting factor in this regard and that the additional thin filament formation that accompanies contractile activation would not have a major effect. More studies are needed to clarify this issue.
The present finding of 20–30% increase in thin filament density is greater than that estimated in other studies (23). We have no clear explanation for the discrepancy. There are a number of factors that could contribute to the variation. Smooth muscle preparations are known to have resting tone, and the tone varies from preparation to preparation. If the resting tone is high, the measured increase in thin filament polymerization due to contractile activation may be underestimated because of the high starting level of polymerized thin filaments. Another source of variation is the alignment of thin filaments with the longitudinal axis of the cell. The filament alignment in the relaxed state may not be as perfect as that in the activated state; this could lead to overestimation of thin filament density increase (due to contractile activation) because a nonperpendicular thin filament found in a relaxed cell cross section may not be recognized as such.
Length-dependent thin filament density. The results presented in Fig. 3 indicate that the thin filament density is higher in muscle cells adapted at a longer length. In the relaxed state, a 50% increase in length resulted in 10.6 ± 1.9% (SE) increase in thin filament density. In the contracted state, the density increase was even greater (20.1 ± 5.8%). It appears that there is synergy between contractile activation and increased strain in augmenting actin polymerization. Statistical analysis (2-way ANOVA), however, revealed that, although the significantly increased (P < 0.05) thin filament formation can be attributed separately to contractile activation and increased cell length, there is no statistically significant interaction between activation and cell length in determining the extent of actin polymerization. Our laboratory has previously shown in the same trachealis preparation that the cell volume is conserved at different cell lengths (17). If the thin filaments (which are short compared with the cell length) are evenly distributed within the cell volume and if there is no polymerization or depolymerization of the filaments, the density of thin filaments at randomly selected cross sections should be the same at different cell lengths. An increase in density, therefore, indicates an increase in the transition of G- to F-actin. Polymerization of myosin filaments was also favored at longer lengths (Fig. 4) as our laboratory has previously found (17). In our previous study, our laboratory also found that both muscle power and shortening velocity increased with adapted length while isometric force was not changed (17). A simple explanation of the finding is that there were additional contractile units (thick filaments) added in series to the contractile apparatus of a cell adapted to a longer length. Because in smooth muscle there are many more thin filaments compared with thick filaments (see Fig. 1), one could argue that there may be enough thin filaments for the additional contractile units needed at longer cell lengths (without additional formation of thin filaments). The present finding indicates that this may not be the case. It appears that, despite a large excess of thin filaments (compared with thick filaments), actin polymerization and depolymerization are still required as part of the process of cell adaptation to different lengths. As discussed above, actin polymerization may be involved in anchoring the cytoskeleton to focal adhesion sites (9), although it is not clear why such polymerization is required.
Thin filament lability vs. thick filament lability.
Lability of myosin filament in smooth muscle has been recognized ever since the ultrastructure of the muscle was examined under electron microscope [see a review by Bagby (3)]. Although it is still controversial as to the extent of lability, it is generally accepted that, compared with those in striated muscle, thick filaments in smooth muscle are structurally less stable. Lability of actin filaments is less well documented. It has been recognized only recently that polymerization and depolymerization associated with the contraction-relaxation cycle in smooth muscle may facilitate plastic adaptation of the muscle to externally applied stress and strain (10, 23). The present results indicate that there was a 63–82% increase in thick filament density due to contractile activation, depending on the adapted length of the muscle cells (Fig. 4). The increase in thin filament density was 20–30% under the same conditions (Fig. 3). It appears, therefore, that the formation of thick filaments is facilitated by contractile activation to a greater extent.
Another difference between thin and thick filaments in terms of their structural stability is shown in the filaments' responses to oscillatory strain. By application of periodic (0.5 Hz) 30% stretches to a relaxed trachealis preparation for 5 min, the thick filament density was found to decrease by 18.4 ± 2.6% (SE), whereas the thin filament density was found to increase by 9.1 ± 1.8 (Fig. 3). Mechanical agitation that caused thick filaments to fall apart apparently had an opposite effect on the thin filaments. In cultured airway smooth muscle cells subjected to long-term oscillatory strain, it has been reported that there was a large increase in the amount of F-actin and that the increase was associated with RhoA activation (29). It is not known whether the increase in thin filament density observed in the present study is related to RhoA activation. It is clear, though, that short-term oscillatory strain did not cause depolymerization of thin filaments, as it did with thick filaments.
The increase in thick filament density due to contractile activation found in the present study was substantially less than that found in one of our previous studies (13). The reason for the discrepancy is not entirely clear. One factor that could explain the difference is the low isometric stress [101.6 ±11.6 (SE) kPa] produced by the muscle strips used in the present study compared with those used in the previous study (161.4 ± 11.3 kPa); the different degree of activation may be one reason for the discrepancy. The oscillatory strain-induced reduction in isometric force found in this study was also significantly less than that found in our laboratory's previous study (18). Again, the reason for the discrepancy is not clear. It is possible that the muscle preparations used in the present study possessed more stray compliance, which reduced the effectiveness of mechanical disruptions on the contractile filaments imposed by the length oscillation. Considering the large variation in results from different studies, it is important that comparisons are made only among muscle preparations dissected from the same trachea and fixed with the same batch of chemicals, as it was done in the present study.
Conclusions. 1) There was regional variation in the density of thin filaments within the cytoplasm of trachealis cells, with the filaments slightly more concentrated near dense bodies. 2) Contractile activation was associated with an increase in the density of thin filaments; the density increase was uniform across the whole cell cross section. 3) Adaptation of trachealis cells to longer lengths resulted in a greater extent of thin filament formation. 4) Application of a brief period of oscillatory strain to the muscle caused a slight increase in thin filament density. 5) Thin filaments were less labile than the thick filaments.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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